Mesoporous catalysts of magnetic nanoparticles and free-radical-producing enzymes, and methods of use

ABSTRACT

A composition comprising mesoporous aggregates of magnetic nanoparticles and free-radical producing enzyme (i.e., enzyme-bound mesoporous aggregates), wherein the mesoporous aggregates of magnetic nanoparticles have mesopores in which the free-radical-producing enzyme is embedded. Methods for synthesizing the enzyme-bound mesoporous aggregates are also described. Processes that use said enzyme-bound mesoporous aggregates for depolymerizing lignin, removing aromatic contaminants from water, and polymerizing monomers polymerizable by a free-radical reaction are also described.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application claims priority from U.S. Provisional Application No.61/568,966, filed Dec. 9, 2011, and U.S. Provisional Application No.61/451,360, filed Mar. 10, 2011, both of which are herein incorporatedby reference in their entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH

This invention was made with government support under contract to theNortheast Sun Grant Initiative at Cornell University US Department ofTransportation Assistance #DTOS59-07-G-00052. The government has certainrights in the invention.

BACKGROUND OF THE DISCLOSURE

Peroxidases (EC 1.11.1) are widely found in biological systems and forma subset of oxidoreductases that reduce hydrogen peroxide (H₂O₂) towater in order to oxidize a large variety of aromatic compounds rangingfrom phenol to aromatic amines. The reaction cycle of peroxidases isquite complex and begins with activation of heme by H₂O₂ to form thetwo-electron activated Compound I (N. C. Veitch, Phytochemistry, 2004,65, 249). Compound I is then reduced by one electron by the oxidation ofthe organic substrate leading to the formation of Compound II that isone electron above the resting state. The second reduction recovers theenzyme to its resting state to start a new cycle. Overall, for eachmolecule of hydrogen peroxide consumed, two aromatic free radicals areproduced and can readily react in secondary reactions.

Peroxidases are highly sensitive to substrate inhibition, mostly byH₂O₂, which can lead to the formation of the reversible inactivated formof the enzyme (Compound III). Their activities are also deterred byproduct inhibition. Therefore, the complex kinetics associated withperoxidase enzymes can restrict their use in many processes andbioprocesses. Increasing the activities of this family of enzymes andtheir tolerance to different process conditions could improve theircurrent use, as well as pave the way for their use in new applications.

BRIEF SUMMARY OF THE DISCLOSURE

It has been discovered herein that bionanocatalysts (BNCs) consisting ofa free-radical-producing (FRP) enzyme, e.g., horseradish peroxidase(HRP), self-assembled with magnetic nanoparticles (MNPs) enhanceenzymatic activity. In particular, it has herein been surprisingly foundthat the self-assembled clusters of FRP enzyme and magneticnanoparticles generally possess faster turnover and lower inhibition ofthe enzyme as compared with the free enzyme or the magnetic nanoparticleclusters without enzyme. It has herein furthermore been found that thesize and magnetization of the MNPs affect the formation and ultimatelythe structure of the BNCs, all of which have a significant impact on theactivity of the entrapped enzymes. Particularly by virtue of theirsurprising resilience under various reaction conditions, the BNCsdescribed herein can be used as an improved FRP agent where other suchagents are currently used, and they can furthermore be used in otherapplications where FRP enzyme has not yet been considered or foundapplicable.

The approach described herein sharply differs from classical methodsthat rely on protein conjugation on surface-modified particles bycomplex biochemistries, oftentimes at the expense of enzymaticactivities and reaction efficiencies. By the instant methodology, HRPkinetics are substantially modified only when the enzymes are in closeassociation with the MNPs, e.g., as a self-assembled cluster(agglomeration) of primary MNP crystallites and peroxidase enzyme. Theoverall activities of the resulting BNCs can advantageously be orders ofmagnitude higher than those of free enzymes or MNPs at biologicallyrelevant substrate concentration.

In one aspect, the invention is directed to a composition in which FRPenzyme is embedded (i.e., entrapped) in magnetic nanoparticles orclusters thereof. In particular embodiments, the composition is amesoporous clustered assembly of magnetic nanoparticles and one or acombination of FRP enzyme. The mesoporous clustered assemblies possessmesopores in which FRP enzyme is embedded. In other embodiments, theforegoing cluster composition includes magnetic nanoparticles that aresurface-coated with gold. In yet other embodiments, the foregoingcluster composition further includes micron- or submicron-sized magneticmicroparticles on which FRP-embedded magnetic nanoparticles reside.

In other aspects, the invention is directed to processes in which theabove-described FRP-embedded magnetic nanoparticle compositions areuseful. In particular embodiments, the FRP-embedded magneticnanoparticle compositions are directed to a process for depolymerizinglignin, a process for removing aromatic contaminants from water, and aprocess for producing a polymer by polymerizing a monomer by a freeradical mechanism.

In yet another aspect, the invention is directed to a process forproducing the FRP-embedded magnetic nanoparticle compositions describedabove. In some embodiments, magnetic nanoparticles or aggregates thereofare first prepared, and FRP enzyme is subsequently absorbed therein orattached thereto. In other embodiments, the FRP-embedded magneticnanoparticle composition is produced by performing a magneticnanoparticle synthesis in the presence of a FRP enzyme, therebyembedding the FRP enzyme in clusters of MNPs by a self-assemblymechanism.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1. X-ray diffraction patterns of magnetite nanoparticlessynthesized at 25° C. and 90° C. The synthesized nanoparticles show thecharacteristic X-ray diffraction pattern of magnetite. The intensity ofthe crystallite diffraction peaks differs between the nanoparticlesformed at 25° C. (smaller size) and the ones formed at 90° C. (biggersize).

FIGS. 2A, 2B. Particle size distribution plots for magnetitenanoparticles synthesized at 25° C. and 90° C. The nanoparticlessynthesized at synthesized at 25° C. and 90° C. have different particlesize distributions. M25 nanoparticles (FIG. 2A) have an average size of8 nm while the average size of M90 nanoparticles (FIG. 2B) is around 10nm.

FIG. 3. Magnetization plot for magnetite nanoparticles synthesized at25° C. and 90° C. The nanoparticles synthesized at 25° C. and 90° C.have a saturated magnetization (M_(s)) of about 20 and 50 emu g⁻¹respectively. Both have negligible remanent magnetization (M_(r)) makingthem superparamagnetic. Their superparamagnetic properties allow forfairly monodispersed magnetic nanoparticles in solution aftersonication.

FIGS. 4A-4C. Particle size distribution plot for M25 and M25-BNC (FIG.4A) and micrographs of M25 (FIG. 4B) and M25-BNC (FIG. 4C). The size ofthe M25 nanoparticle aggregates increases when the enzyme (HRP) is addedto the initially monodispersed magnetic nanoparticles. The presence ofthe enzyme increases the overall diameter of the clusters.

FIGS. 5A-5G. Micrograph of M90 nanoparticle clusters (FIG. 5A) andcorresponding size plots (FIGS. 5B, 5C), micrograph of M90-BNCnanoparticle clusters (FIG. 5D) and corresponding size plots (FIGS. 5E,5F), and size distribution plot comparing M90 and M90-BNC clusters (FIG.5G).

FIG. 6. Graph plotting the isotherms of the quantity of entrapped enzymeper quantity of enzyme in solution per available surface area of MNPs.The quantity of enzymes entrapped in the clusters (standardized persurface area of the MNPs) was measured for M90 and M25. The aggregateformed with M90 MNPs contains more enzyme than the one formed with M25MNPs for an initial amount of enzyme in solution. Below 10 nmol·m⁻² ofenzyme in solution, 100% of the enzymes are entrapped in the clustersformed with M90, while about 50% are trapped in the M25 clusters.

FIG. 7. Pore size distribution plot for M25 magnetic nanoparticles andM25-BNC enzyme clusters. M25-BNCs are mesoporous with pores below 50 nmin diameter. Smaller nanoparticles form aggregates with smaller poresize. The pore size distribution is affected by the presence of theenzyme, and the total pore volume is lower than the total pore volume ofthe M25 MNPs clusters, thus indicating that the enzymes occupy themesoporous space.

FIG. 8. Pore size distribution plot for M90 magnetic nanoparticles andM90-BNC enzyme clusters. The cluster formed by the aggregation of M90MNPs are mesoporous, with pores below 50 nm in diameter. Largernanoparticles form larger aggregates with larger pore size. The poresize distribution is affected by the presence of the enzyme, and thetotal pore volume is lower than the total pore volume of the M90 MNPsclusters, thus indicating that the enzymes occupy the mesoporous space.

FIGS. 9A, 9B. Graphs plotting activity (Phenol AAP assay, in U·mmole⁻¹)as a function of incubation time for M25-BNC (FIG. 9A) and for M90-BNC(FIG. 9B) at 4 μg/mL and 8 μg/mL concentrations to determine the effectof incubation time on the activity of the BNC.

FIG. 10. Graph plotting normalized activity as a function ofconcentration of magnetic nanoparticles (HRP-BNC) used in the assay (inμg/mL) to determine effect of incubation time on the activity of theHRP-BNC. P5 protocol: 3% (vol/vol) premix from diluted 1:1 (vol/vol)solutions of HRP+nanoparticles (M90); P1 protocol: 3% (vol/vol) premixfrom high concentration stock solution of HRP and M90; P3 protocol: 14%(vol/vol) premix 1:1 (vol/vol) suspensions of HRP+nanoparticles; P4protocol: no premix, add enzymes and particles separately in the assaymix. Protocols P5 and P1 are comparing the way to mix the enzyme and thenanoparticles (M90) to form BNCs. The effect of BNC concentration in theassay is shown by P3 and P5. For comparison, P4 shows the effect of noincubation at all. The self-assembly depends on the concentration ofMNPs in the premix (enzyme+MNPs). High concentration of MNPs leads tolower activities due to over-agglomeration.

FIGS. 11A, 11B. 96-well plate for the phenol/AAP assay and lay-out usedfor determining effect of PBS buffer on BNC formation and activity (FIG.11A), and chart showing increased activity data (normalized by theactivity of the free enzyme) for M90-BNCs in H₂O and varyingconcentration of phosphate buffer saline (PBS) (FIG. 11B). Theconditions to form BNCs were investigated with the phenol/AAP assay witha fixed concentration of H₂O₂ (i.e., peroxide) at 1 mM. BNCs were formedin PBS buffer at different ionic strengths (up to 200 mM). The increasein activity decreased when the ionic strength of the buffer increased.The highest increase in activity was observed for the BNCs formed inwater (no ionic compensation charge). BNCs formed in water showed anincrease in activity up to 26 times and can be used at higherconcentration of PBS in the assay.

FIGS. 12A, 12B. 96-well plate for the phenol/AAP assay and lay-out usedfor determining effect of malonate buffer on BNC formation and activity(FIG. 12A), and chart showing increased activity data (normalized by theactivity of the free enzyme) for M90-BNCs in H₂O and varyingconcentration of sodium malonate buffer (SMB) (FIG. 12B). The conditionsto form BNCs were investigated with the phenol/AAP assay with a fixedconcentration of peroxide (1 mM). BNCs were formed in an organic buffer(sodium malonate: SMB) at different ionic strengths (up to 200 mM). Theincrease in activity decreased when the ionic strength of the bufferincreased, but was an order of magnitude higher than the BNCs formed inPBS. The highest increase in activity was observed for the BNCs formedin water (no ionic compensation charge). BNCs formed in SMB showed anincrease in activity up to 26 times and can be used at higherconcentration of SMB in the assay.

FIGS. 13A, 13B. 96-well plate for the phenol/AAP assay and lay-out usedfor determining effect of tartrate buffer on BNC formation and activity(FIG. 13A), and chart showing increased activity data (normalized by theactivity of the free enzyme) for M90-BNCs in H₂O and varyingconcentration of sodium tartrate buffer (STB) (FIG. 13B). The conditionsto form BNCs were investigated with the phenol/AAP assay with a fixedconcentration of peroxide (1 mM). BNCs were formed in an organic buffer(sodium tartrate: STB) at different ionic strengths (up to 200 mM). Theincrease in activity decreased when the ionic strength of the bufferincreased, but was an order of magnitude higher than the BNCs formed inPBS. The highest increase in activity was observed for the BNCs formedin water (no ionic compensation charge). BNCs formed in STB showed anincrease in activity up to 24 times and can be used at higherconcentration of STB in the assay.

FIG. 14. Graph plotting molecular activity (Phenol/AAP assay, inU·mmole⁻¹) as a function of peroxide concentration for HRP-BNCnanoparticle aggregates to determine effect of enzyme saturation of themesoporous spaces. Surprisingly, the MNP aggregates saturated withenzyme did not increase the enzyme activity, whereas BNCs with anon-saturated amount of enzyme showed increased activities.

FIG. 15. Graph plotting K_(m) kinetic constant as a function ofnanoparticle concentration for free HRP, M25 BNC and M90 BNC in order todetermine substrate affinity constant for free HRP and BNCs formed withM25 and M90. The K_(m) kinetic constant estimated was by fitting thevelocities plots of the HRP-BNCs. The affinity constant was increased inthe case of BNCs formed with M25, thus indicating a better utilizationof peroxide. No difference in K_(m) was observed in the case of enzymesentrapped in the mesoporous space of the BNCs formed with M90.

FIG. 16. Graph plotting V_(max) as a function of nanoparticleconcentration for free HRP, M25 BNC and M90 BNC in order to determinemaximal velocity constant for free HRP and BNCs formed with M25 and M90.The V_(max) kinetic constant was estimated by fitting the velocitiesplots of the HRP-BNCs. The maximal velocity was increased in the case ofBNCs formed with M25 compared to the V_(max) of the free enzymes, thusindicating a better utilization of peroxide. No difference in V_(max)was observed in the case of the enzymes entrapped in the mesoporousspace of the BNCs formed with M90 and their free counterpart.

FIG. 17. Graph plotting K_(i) kinetic constant as a function ofnanoparticle concentration for free HRP, M25 BNC and M90 BNC in order todetermine inhibition constants for free HRP and BNCs formed with M25 andM90. The K_(i) kinetic constant was estimated by fitting the velocitiesplots of the BNCs made with HRP. The substrate inhibition constant wasdrastically increased in the case of BNCs formed with M90, thusindicating a better protection of the enzyme against inhibition whenentrapped in the mesoporous space of the BNCs. Inhibition of M25-BNC wasin the same range, if not lower, than for the free enzymes.

FIG. 18. Graph plotting k_(cat) kinetic constant as a function ofnanoparticle concentration for free HRP, M25 BNC and M90 BNC in order todetermine turnover constant for free HRP and BNCs formed with M25 andM90. The k_(cat) kinetic constant was estimated by fitting thevelocities plots of the BNCs made with HRP. The turnover constant wasdrastically increased in the case of BNCs formed with M25 compared tothe free enzyme, thus indicating a better utilization of the substrates.The k_(cat) of M90-BNCs was also increased for lower concentrations ofenzymes.

FIGS. 19A, 19B. Graphs plotting normalized activity (phenol/AAP assay)as a function of assay buffer and pH (FIG. 19A) and graphs plottingvelocity as a function of assay buffer in pH (FIG. 19B). BNCs wereformed with HRP (1 nM) and M90 MNPs (4 μg/ml) in water, and were testedat different pH conditions with the phenol/AAP assay at 5 mM peroxideand 5 mM buffers. The increase in activity was observed for all buffersand at all pHs.

FIGS. 20A, 20B. Graphs plotting normalized activity as a function ofassay buffer and pH (FIG. 20A) and graphs plotting velocity as afunction of assay buffer in pH (FIG. 20B) for the phenol/AAP assay. BNCswere formed with HRP (1 nM) and M90 MNPs (4 μg/ml) in water, and weretested at different temperature conditions with the phenol/AAP assay at5 mM peroxide and 3 mM buffers. The increase in activity was observedfor all buffers and at all temperatures, and was maximal at 25° C. Theobserved increase compared to the free enzyme appears to be the resultof the lower velocity of the free enzyme, thus indicating that the BNCsare more efficient at lower temperature.

FIG. 21. Graph plotting molecular activity (phenol/AAP assay inU·mmole⁻¹) as a function of peroxide concentration (M) for BNCs formedwith versatile peroxidase and M90 and functionalized M90. The BNCs wereformed with Versatile Peroxidase (VeP: commercial form, no additionalpurification) and assayed with the Phenol/AAP (no manganese). M90-MNPswere functionalized with organic polymers to modify the charge of thesurfaces. All BNCs showed increased activity compared to the free VeP.The maximal activity was observed for non-modified BNCs.

FIG. 22. Velocities plot of BNCs formed with Manganese Peroxidase andgold-coated MNPs compared with M90-MNPs. BNCs were formed with M90 orgold-coated M90 and Manganese Peroxidase (FLPC anion exchange purifiedform, 5 nM). In this assay, the Manganese Peroxidase produces Mn³⁺reactive cations, which complex with organic acids and oxidizesdimethoxyphenol (colorimetric reagent). The initial rate of the reactionwas doubled in the case of BNCs formed with M90 compared to the freeenzyme. The velocity of the reaction was multiplied by a factor of fourwith gold-coated BNCs compared to the free enzyme.

FIG. 23. Graph plotting increased activities of laccase BNCs (Lac-BNCs,normalized activity) as a function of buffer concentration and pH. Theassay (phenol/AAP without peroxide) was conducted in different buffersat different pHs. The optimal pH for laccase enzymes is around 5. Theincrease in activity (up to 60%) was only observed for low and high pHand was higher for high molarity buffers.

FIGS. 24A-24D. Graph plotting the kinetics of Homovanillic Acidfluorescent dimer formation to determine increased velocities ofperoxidase using mixed enzyme BNCs, i.e., clusters with HorseradishPeroxidase (HRP) in the center (core) and Glucose Oxidase (Gox) in theouter layers (shell), clusters with Gox in the center and HRP at theouter layers, or random distribution. The BNCs were formed withgold-coated MNPs and a ratio of Gox to HRP of 1. The HVA assay was usedwith increasing concentration of glucose. In this system, the GlucoseOxidase converts the glucose to peroxide and gluconolactone and the HRPuses the peroxide to polymerize the homovanillic acid to its fluorescentdimer. The kinetics show that the distribution of the enzymes in themesoporous space matters.

FIG. 25. Velocity plot of a mixed enzyme gold-coated BNC (HVA assay,Glucose Oxidase:HRP in a 1:1 ratio). In this configuration, the velocity(initial rate of the reaction) were higher for the Gox/HRP BNCs in thelower range of glucose compared to the free enzymes. In the higherglucose range, BNCs activity was similar to the one of the free enzyme.No substrate inhibition was observed for this ratio of Gox/HRP andglucose concentration range. The initial rate of the reaction wasdoubled for the HRP core BNCs compared to the randomly distributed ones.These results demonstrate that the polyenzyme systems can be used, andthat the distribution of the enzymes can be controlled to favorablyaffect the activities of the system.

FIGS. 26A, 26B. SEM micrographs of commercial magnetic microparticles(FIG. 26A) and the microparticles surface-functionalized withgold-coated BNCs (Glucose Oxidase and HRP). The preformed BNCs can beimmobilized on ferromagnetic microparticles with a remanentmagnetization. This new material possesses the advantages of having theenzymes entrapped in the mesoporous space of the magnetic clusters andthe additional advantage of having them immobilized on a stable andhighly magnetic material.

FIG. 27. Graph plotting the kinetics of Homovanillic Acid fluorescentdimer formation produced by Glucose Oxidase/HRP μBNCs. The μBNCs weremade using gold-coated Gox/HRP BNCs with clusters with HRP in the centerand Gox in the outer layers, clusters with Gox in the center and HRP atthe outer layers, or random distribution. The velocity of the core/shellμBNCs was higher than the free enzyme or the randomly distributed BNCs.These results demonstrate that the BNCs can be immobilized on themicroparticles and maintain the increase in activity observed for theBNCs alone. Gox:HRP=1:1. Enzyme:MNPs=1:2. MNPs:microsphere=1:100 (W/W).

FIGS. 28A-28C. Photographs demonstrating the superior ability of μBNCsto be captured by an external magnet. FIG. 28A compares solutions ofHRP/Gox (left) and HRP/Gox BNCs (right) in 1 mM glucose solution. FIG.28B compares the same solutions after exposure to an external magnet.FIG. 28C shows the μBNCs stirred by an external rotating magnet. Asshown, the μBNCs can be retrieved from solution after the reaction in amatter of seconds even with low magnetic fields, and can also be easilystirred with external magnetic fields. The darker color of theHRP/Gox-BNC system immobilized on microparticles (right tube) alsoindicated that the extent of the reaction is higher compared to the freeenzyme (left tube) after 10 hours.

FIG. 29. Wavelength absorbance spectra for HRP-BNC, LiP-BNC, VeP-BNC(without Mn), and MnP-BNC using M90 magnetite nanoparticles. The spectrafrom the free enzyme at the same concentration were subtracted. Thefungal peroxidases entrapped in the mesoporous space of M90-BNCs showhigher activities relative their free counterpart (i.e., increasedrelease of aromatics from lignin depolymerization).

FIG. 30. Chart plotting the increase in soluble aromatic molecules fromKraft lignin with free lignin peroxidase (neat LiP), and LiP+M8 (BNCformed with 2 nM LiP and M90 at 8 μg/ml), and LiP+M16 (BNC formed with 2nM LiP+M90 at 16 μg/ml). The release of soluble aromatics from Kraftlignin by BNCs was observed at characteristic absorbance wavelengths.BNCs were formed with Lignin Peroxidase at 2 nM enzyme (commercial form,not purified). The OD values for supernatant at T0 (initial condition)and the nanoparticle background were subtracted.

FIG. 31. Chart plotting the increase in soluble aromatic molecules fromKraft lignin with free Versatile Peroxidase (neat VP), VP+M8 (BNC formedwith 2 nM VP and M90 at 8 μg/ml), and VP+M16 (BNC formed with 2 nM VPand M90 at 16 μg/ml). The release of soluble aromatics from Kraft ligninby BNCs was observed at characteristic absorbance wavelengths. BNCs wereformed with Versatile Peroxidase at 2 nM enzyme (commercial form, notpurified). The OD values for supernatant at T0 (initial condition) andthe nanoparticle background were subtracted.

FIG. 32. Chart plotting the increase in soluble aromatic molecule fromKraft lignin with free Versatile Peroxidase (neat VP) in presence of Mn,VP+M8 (BNC formed with 2 nM VP and M90 at 8 μg/ml), and VP+M16 (BNCformed with 2 nM VP and M90 at 16 μg/ml). The release of solublearomatics from Kraft lignin by BNCs was observed at characteristicabsorbance wavelengths. BNCs were formed with Versatile Peroxidase at 2nM enzyme (commercial form, not purified). The OD values for supernatantat T0 (initial condition) and the nanoparticle background weresubtracted.

FIG. 33. Chart plotting the increase in soluble aromatic molecules fromKraft lignin with free Manganese Peroxidase (neat MnP), MnP+M8 (BNCformed with 1 nM MnP and M90 at 8 μg/ml), and MnP+M16 (BNC formed with 1nM and M90 at 16 μg/ml). The release of soluble aromatics from Kraftlignin by BNCs was observed at characteristic absorbance wavelengths.BNCs were formed with Manganese Peroxidase at 1 nM enzyme (commercialform, not purified). The OD values for supernatant at T0 (initialcondition) and the nanoparticle background were subtracted.

FIG. 34. Photograph demonstrating phenol polymerization andprecipitation of polyphenols by HRP and BNCs formed with M90. Thecondensed polyphenols were precipitated by NaCl and centrifuged, and theremaining phenol in solution was measured by absorbance. The absorbancewas corrected for the presence of MNPs.

FIG. 35. Graph plotting average phenol removal (%) as a function ofMnP/HRP ratio to determine effect of nanoparticle to enzyme ratio on thepercentage of phenol removal. The total enzyme (horseradish peroxidase)quantity and peroxide and phenol concentrations (1 mM each) were fixed.The proportion of MNPs was increased to form the BNCs. The quantity ofphenol removed from solution increased when the enzyme was entrapped inthe mesoporous space of the magnetic clusters of M90. The efficiency ofphenol removal was close to 95%.

FIG. 36. Graph plotting % phenol removal as a function of the ratio ofperoxide to phenol to determine the effect of hydrogen peroxide tophenol ratio on phenol removal by HRP and BNCs thereof. The phenol finalconcentration was fixed to 1 mM. The quantity of peroxide was varied.The efficiency of phenol removal increased up to 100% for aperoxide/phenol ratio of 1.1, for which the efficiency of the freeenzyme was only around 30%. The efficiency of the free enzyme decreasedwhen the peroxide concentration increased due to substrate inhibition.On the contrary, the BNCs showed little or no inhibition in the rangetested depending on the MNP/HRP ratio.

DETAILED DESCRIPTION OF THE DISCLOSURE

In one aspect, the invention is directed to a free-radical producing(FRP) composition that includes magnetic nanoparticles bound to FRPenzyme. Magnetic nanoparticles bound to FRP enzyme is herein alsoreferred to as a “bionanocatalyst” or “BNC”. As used herein, the term“bound” is intended to include any of the means by which FRP enzyme canbe attached to magnetic nanoparticles without the release of FRP enzymefrom the magnetic nanoparticles under conditions in which they are usedor stored for later use. The FRP enzyme can be bound by, for example,covalent, ionic, hydrogen bonding, affinity, or van der Waalsinteractions. The FRP enzyme may be located anywhere on the magneticnanoparticle, e.g., on the surface and/or embedded within the magneticnanoparticle, such as in mesopores of the magnetic nanoparticles if themagnetic nanoparticles are porous. As used herein, the term “magnetic”encompasses all types of useful magnetic characteristics, includingpermanent magnetic, superparamagnetic, paramagnetic, ferromagnetic, andferrimagnetic behaviors.

The magnetic nanoparticle or BNC has a size in the nanoregime, i.e.,generally no more than 500 nm. As used herein, the term “size” can referto a diameter of the magnetic nanoparticle when the magneticnanoparticle is approximately or substantially spherical. In a casewhere the magnetic nanoparticle is not approximately or substantiallyspherical (e.g., substantially ovoid or irregular), the term “size” canrefer to either the longest the dimension or an average of the threedimensions of the magnetic nanoparticle. The term “size” may also referto an average of sizes over a population of magnetic nanoparticles(i.e., “average size”). In different embodiments, the magneticnanoparticle has a size of no more than, for example, 500 nm, 400 nm,300 nm, 200 nm, 100 nm, 50 nm, 40 nm, 30 nm, 25 nm, 20 nm, 15 nm, 10 nm,5 nm, 4 nm, 3 nm, 2 nm, or 1 nm, or a size within a range bounded by anytwo of the foregoing exemplary sizes.

The magnetic nanoparticles described above or BNCs thereof may beclustered, i.e., as aggregates or agglomerations, in which case theabove-described magnetic nanoparticles are considered to be primarynanoparticles (i.e., primary crystallites) and the magnetic nanoparticlesizes provided above can be considered to be primary nanoparticle sizes.The aggregates generally have a size (i.e., secondary size) of at least5 nm. In different embodiments, the aggregates have a size of precisely,about, or at least, for example, 5 nm, 8 nm, 10 nm, 12 nm, 15 nm, 20 nm,25 nm, 30 nm, 35 nm, 40 nm, 45 nm, 50 nm, 60 nm, 70 nm, 80 nm, 90 nm,100 nm, 150 nm, 200 nm, 300 nm, 400 nm, 500 nm, 600 nm, 700 nm, or 800nm, or a size within a range bounded by any two of the foregoingexemplary sizes.

Typically, the primary and/or aggregate magnetic nanoparticles or BNCsthereof have a distribution of sizes, i.e., they are generally dispersedin size, either narrowly or broadly dispersed. In different embodiments,any range of primary or aggregate sizes can constitute a major or minorproportion of the total range of primary or aggregate sizes. Forexample, in some embodiments, a particular range of primary particlesizes (for example, at least 1, 2, 3, 5, or 10 nm and up to 15, 20, 25,30, 35, 40, 45, or 50 nm) or a particular range of aggregate particlesizes (for example, at least 5, 10, 15, or 20 nm and up to 50, 100, 150,200, 250, or 300 nm) constitutes at least or above 50%, 60%, 70%, 80%,90%, 95%, 98%, 99%, or 100% of the total range of primary particlesizes. In other embodiments, a particular range of primary particlesizes (for example, less than 1, 2, 3, 5, or 10 nm, or above 15, 20, 25,30, 35, 40, 45, or 50 nm) or a particular range of aggregate particlesizes (for example, less than 20, 10, or 5 nm, or above 25, 50, 100,150, 200, 250, or 300 nm) constitutes no more than or less than 50%,40%, 30%, 20%, 10%, 5%, 2%, 1%, 0.5%, or 0.1% of the total range ofprimary particle sizes.

The aggregates of magnetic nanoparticles (i.e., “aggregates”) or BNCsthereof can have any degree of porosity, including a substantial lack ofporosity depending upon the quantity of individual primary crystallitesthey are made of. In particular embodiments, the aggregates aremesoporous by containing interstitial mesopores (i.e., mesopores locatedbetween primary magnetic nanoparticles, formed by packing arrangements).The mesopores are generally at least 2 nm and up to 50 nm in size. Indifferent embodiments, the mesopores can have a pore size of preciselyor about, for example, 2, 3, 4, 5, 10, 12, 15, 20, 25, 30, 35, 40, 45,or 50 nm, or a pore size within a range bounded by any two of theforegoing exemplary pore sizes. Similar to the case of particle sizes,the mesopores typically have a distribution of sizes, i.e., they aregenerally dispersed in size, either narrowly or broadly dispersed. Indifferent embodiments, any range of mesopore sizes can constitute amajor or minor proportion of the total range of mesopore sizes or of thetotal pore volume. For example, in some embodiments, a particular rangeof mesopore sizes (for example, at least 2, 3, or 5, and up to 8, 10,15, 20, 25, or 30 nm) constitutes at least or above 50%, 60%, 70%, 80%,90%, 95%, 98%, 99%, or 100% of the total range of mesopore sizes or ofthe total pore volume. In other embodiments, a particular range ofmesopore sizes (for example, less than 2, 3, 4, or 5 nm, or above 10,15, 20, 25, 30, 35, 40, 45, or 50 nm) constitutes no more than or lessthan 50%, 40%, 30%, 20%, 10%, 5%, 2%, 1%, 0.5%, or 0.1% of the totalrange of mesopore sizes or of the total pore volume.

The magnetic nanoparticles can have any of the compositions known in theart. In some embodiments, the magnetic nanoparticles are or include azerovalent metallic portion that is magnetic. Some examples of suchzerovalent metals include cobalt, nickel, and iron, and their mixturesand alloys. In other embodiments, the magnetic nanoparticles are orinclude an oxide of a magnetic metal, such as an oxide of cobalt,nickel, or iron, or a mixture thereof. In some embodiments, the magneticnanoparticles possess distinct core and surface portions. For example,the magnetic nanoparticles may have a core portion composed of elementaliron, cobalt, or nickel and a surface portion composed of a passivatinglayer, such as a metal oxide or a noble metal coating, such as a layerof gold, platinum, palladium, or silver. In other embodiments, metaloxide magnetic nanoparticles or aggregates thereof are coated with alayer of a noble metal coating. The noble metal coating may, forexample, reduce the number of charges on the magnetic nanoparticlesurface, which may beneficially increase dispersibility in solution andbetter control the size of the BNCs. Noble metal coating protects theparticles against oxidation, solubilization by leaching or by chelationwhen chelating organic acids such as citrate, malonate, tartrate forexamples are used in the biochemical reactions or processes. Thepassivating layer can have any suitable thickness, and particularly, atleast, up to, or less than, for example, 0.1 nm, 0.2 nm, 0.3 nm, 0.4 nm,0.5 nm, 0.6 nm, 0.7 nm, 0.8 nm, 0.9 nm, 1 nm, 2 nm, 3 nm, 4 nm, 5 nm, 6nm, 7 nm, 8 nm, 9 nm, or 10 nm, or a thickness in a range bounded by anytwo of these values.

In particular embodiments, the magnetic nanoparticles have an iron oxidecomposition. The iron oxide composition can be any of the magnetic orsuperparamagnetic iron oxide compositions known in the art, e.g.,magnetite (Fe₃O₄), hematite (α-Fe₂O₃), maghemite (γ-Fe₂O₃), or a spinelferrite according to the formula AB₂O₄, wherein A is a divalent metal(e.g., Zn²⁺, Ni²⁺, Mn²⁺, Co²⁺, Ba²⁺, Sr²⁺, or combination thereof) and Bis a trivalent metal (e.g., Fe³⁺, Cr³⁺, or combination thereof).

In some embodiments, the magnetic nanoparticles or aggregates thereof orBNCs thereof reside on the surface of ferromagnetic sub-micrometricparticles. By residing on the surface of ferromagnetic microparticles,the magnetic nanoparticles or aggregates or BNCs are attached to thesurface of the ferromagnetic microparticles by any suitable associative,adsorptive, or bonding interaction. The ferromagnetic microparticles mayor may not be coated with a metal oxide or noble metal coating layer.Moreover, the ferromagnetic microparticles may possess any suitablesurface groups, as well known in the art, which may promote attachmentof the magnetic nanoparticles thereto. In different embodiments, theferromagnetic microparticles have a size of about, precisely, or atleast 20, 30, 40, or 50, 60, 70, 80, 90, 100, 100, 200, 300, 400, 500,600, 700, 800, 900, 1000 nm, or a size within a range bounded by any twoof the foregoing exemplary sizes. By virtue of their larger size(ultrastructure), BNCs attached onto the surface of ferromagneticparticles can be more easily captured by an external magnetic field. Thelarger size also helps to preserve enzymatic activities. These biggermagnetic particles can be easily captured by external magnetic fields.BNCs attached onto the surface of ferromagnetic submicrometric particlesare not prone to over aggregation when subjected to magnetic fields.

The magnetic nanoparticles or aggregates thereof or BNCs thereof possessany suitable degree of magnetism. For example, the magneticnanoparticles or aggregates thereof or BNCs thereof can possess asaturated magnetization (M_(s)) of at least or up to 5, 10, 15, 20, 25,30, 40, 45, 50, 60, 70, 80, 90, or 100 emu/g. The magnetic nanoparticlesor aggregates thereof preferably possess a remanent magnetization(M_(r)) of no more than (i.e., up to) or less than 5 emu/g, and morepreferably, up to or less than 4 emu/g, 3 emu/g, 2 emu/g, 1 emu/g, 0.5emu/g, or 0.1 emu/g. The surface magnetic field of the magneticnanoparticles or aggregates thereof can be about or at least, forexample, 5, 10, 50, 100, 200, 300, 400, 500, 600, 700, 800, 900, or 1000Gauss (G), or a magnetic field within a range bounded by any two of theforegoing values.

The magnetic nanoparticles or aggregates thereof can be made to adsorb asuitable amount of FRP enzyme, up to or below a saturation level,depending on the application, to produce the resulting BNC. In differentembodiments, the magnetic nanoparticles or aggregates thereof may adsorbabout, at least, up to, or less than, for example, 1, 5, 10, 15, 20, 25,or 30 pmol/m² of FRP enzyme. Alternatively, the magnetic nanoparticlesor aggregates thereof may adsorb an amount of FRP enzyme that is about,at least, up to, or less than, for example, 10%, 20%, 30%, 40%, 50%,60%, 70%, 80%, 90%, or 100% of a saturation level.

The magnetic nanoparticles or aggregates thereof or BNCs thereof possessany suitable pore volume. For example, the magnetic nanoparticles oraggregates thereof can have a pore volume of about, at least, up to, orless than, for example, 0.01, 0.05, 0.1, 0.15, 0.2, 0.25, 0.3, 0.35,0.4, 0.45, 0.5, 0.55, 0.6, 0.65, 0.7, 0.75, 0.8, 0.85, 0.9, 0.95, or 1cm³/g, or a pore volume within a range bounded by any two of theforegoing values.

The magnetic nanoparticles or aggregates thereof or BNCs thereof possessany suitable specific surface area. For example, the magneticnanoparticles or aggregates thereof can have a specific surface area ofabout, at least, up to, or less than, for example, 50, 60, 70, 80, 90,100, 110, 120, 130, 140, 150, 160, 170, 180, 190, or 200 m²/g.

The FRP enzyme can be any enzyme that produces free radicals. Moreover,the FRP enzyme can be from any source, e.g., fungal, microbial, animal,or plant. In particular embodiments, the FRP enzyme is an oxidoreductasebelonging to the EC 1 family of enzymes. The EC 1 oxidoreductase can be,for example, an EC 1.1 oxidoreductase acting on the CH—OH groups ofdonors, an EC 1.2 oxidoreductase acting on the aldehyde or oxo group ofdonors, an EC 1.3 oxidoreductase acting on the CH—CH group of donors, anEC 1.4 oxidoreductase acting on the CH—NH₂ group of donors, an EC 1.5oxidoreductase acting on the CH—NH group of donors, an EC 1.6oxidoreductase acting on NADH or NADPH, an EC 1.7 oxidoreductase actingon various nitrogenous compounds as donors, an EC 1.8 oxidoreductaseacting on a sulfur group as donor, an EC 1.9 oxidoreductase acting on aheme group of donors, an EC 1.10 oxidoreductase acting on diphenols andrelated substances as donors, an EC 1.11 oxidoreductase acting onperoxide as an acceptor, an EC 1.12 oxidoreductase acting on hydrogen asa donor, an EC 1.13 oxidoreductase acting on single donors withincorporation of molecular oxygen (oxygenases), an EC 1.14oxidoreductase acting on paired donors with incorporation or reductionof molecular oxygen, an EC 1.15 oxidoreductase acting on superoxide asan acceptor, an EC 1.16 oxidoreductase that oxidize metal ions, an EC1.17 oxidoreductase acting on CH or CH₂ groups, an EC 1.18oxidoreductase acting on iron-sulfur proteins as a donor, an EC 1.19oxidoreductase acting on reduced flavodoxin as a donor, an EC 1.20oxidoreductase acting on phosphorus or arsenic as a donor, an EC 1.21oxidoreductase acting on X—H and Y—H to form an X—Y bond, an EC 1.97oxidoreductase, an EC 1.98 oxidoreductase that uses hydrogen as areductant, and an EC 1.99 oxidoreductase that uses oxygen as an oxidant.The oxidoreductase may also be more particularly identified as belongingto a sub-genus of any of the EC 1.1 groupings provided above.

In a first particular set of embodiments, the FRP enzyme is selectedfrom the EC 1.1 genus of oxidoreductase enzymes. The EC 1.1 enzyme canfurther be identified as belonging to any of the following sub-genuses:EC 1.1.1 with NAD or NADP as acceptor, EC 1.1.2 with a cytochrome asacceptor, EC 1.1.3 with oxygen as acceptor, EC 1.1.4 with disulfide asacceptor, EC 1.1.5 with quinone or similar compound as acceptor, and EC1.1.99 with other acceptors. In more particular embodiments, the FRPenzyme is identified as belonging to a sub-genus of any of the EC 1.1sub-genuses provided above. For example, the FRP enzyme can beidentified as belonging to any of the sub-genuses of EC 1.1.3, such asEC 1.1.3.3 (malate oxidase), EC 1.1.3.4 (glucose oxidase), EC 1.1.3.5(hexose oxidase), EC 1.1.3.6 (cholesterol oxidase), EC 1.1.3.7(aryl-alcohol oxidase), EC 1.1.3.8 (L-gulonolactone oxidase), EC 1.1.3.9(galactose oxidase), EC 1.1.3.10 (pyranose oxidase), EC 1.1.3.11(L-sorbose oxidase), EC 1.1.3.12 (pyridoxine 4-oxidase), EC 1.1.3.13(alcohol oxidase), EC 1.1.3.14 (catechol oxidase), EC 1.1.3.15(2-hydroxy acid oxidase), EC 1.1.3.16 (ecdysone oxidase), EC 1.1.3.17(choline oxidase), EC 1.1.3.18 (secondary-alcohol oxidase), EC 1.1.3.19(4-hydroxymandelate oxidase), EC 1.1.3.20 (long-chain alcohol oxidase),EC 1.1.3.21 (glycerol-3-phosphate oxidase), EC 1.1.3.22, EC 1.1.3.23(thiamine oxidase), EC 1.1.3.24 (L-galactonolactone oxidase), EC1.1.3.25, EC 1.1.3.26, EC 1.1.3.27 (hydroxyphytanate oxidase), EC1.1.3.28 (nucleoside oxidase), EC 1.1.3.29 (N-acylhexosamine oxidase),EC 1.1.3.30 (polyvinyl alcohol oxidase), EC 1.1.3.31, EC 1.1.3.32, EC1.1.3.33, EC 1.1.3.34, EC 1.1.3.35, EC 1.1.3.36, EC 1.1.3.37D-arabinono-1,4-lactone oxidase), EC 1.1.3.38 (vanillyl alcoholoxidase), EC 1.1.3.39 (nucleoside oxidase, H₂O₂ forming), EC 1.1.3.40(D-mannitol oxidase), and EC 1.1.3.41 (xylitol oxidase).

In a second particular set of embodiments, the FRP enzyme is selectedfrom the EC 1.10 genus of oxidoreductase enzymes. The EC 1.10 enzyme canfurther be identified as belonging to any of the following sub-genuses:EC 1.10.1 with NAD or NADP as acceptor EC 1.10.2 with cytochrome asacceptor, EC 1.10.3 with oxygen as acceptor, and EC 1.10.99 with otheracceptors. The EC 1.10.1 enzyme can be more specifically, for example,EC 1.10.1.1, i.e., trans-acenaphthene-1,2-diol dehydrogenase. The EC1.10.2 enzyme can be more specifically, for example, EC 1.10.2.1(cytochrome-b5 reductase) or EC 1.10.2.2 (cytochrome-c reductase). TheEC 1.10.3 enzyme can be more specifically, for example, EC 1.10.3.1(catechol oxidase), EC 1.10.3.2 (laccase), EC 1.10.3.3 (L-ascorbateoxidase), EC 1.10.3.4 (o-aminophenol oxidase), EC 1.10.3.5(3-hydroxyanthranilate oxidase), EC 1.10.3.6 (rifamycin-B oxidase), EC1.10.3.7, or EC 1.10.3.8. The EC 1.10.99 enzyme can be morespecifically, for example, EC 1.10.99.1 (plastoquinol-plastocyaninreductase), EC 1.10.99.2 (ribosyldihydronicotinamide dehydrogenase,quinone), or EC 1.10.99.3 (violaxanthin de-epoxidase).

In a third particular set of embodiments, the FRP enzyme is selectedfrom the EC 1.11 genus of oxidoreductase enzymes. The EC 1.11 enzyme canfurther be identified as belonging to the sub-genus EC 1.11.1(peroxidases). The EC 1.11.1 enzyme can be more specifically, forexample, EC 1.11.1.1 (NADH peroxidase), EC 1.11.1.2 (NADPH peroxidase),EC 1.11.1.3 (fatty acid peroxidase), EC 1.11.1.4, EC 1.11.1.5(cytochrome-c peroxidase), EC 1.11.1.6 (catalase), EC 1.11.1.7(peroxidase), EC 1.11.1.8 (iodide peroxidase), EC 1.11.1.9 (glutathioneperoxidase), EC 1.11.1.10 (chloride peroxidase), EC 1.11.1.11(L-ascorbate peroxidase), EC 1.11.1.12 (phospholipid-hydroperoxideglutathione peroxidase), EC 1.11.1.13 (manganese peroxidase), EC1.11.1.14 (diarylpropane peroxidase), or EC 1.11.1.15 (peroxiredoxin).

In particular embodiments, the FRP enzyme is a peroxidase. Theperoxidase may also be further specified by function, e.g., a ligninperoxidase, manganese peroxidase, or versatile peroxidase. Theperoxidase may also be specified as a fungal, microbial, animal, orplant peroxidase. The peroxidase may also be specified as a class I,class II, or class III peroxidase. The peroxidase may also be specifiedas a myeloperoxidase (MPO), eosinophil peroxidase (EPO), lactoperoxidase(LPO), thyroid peroxidase (TPO), prostaglandin H synthase (PGHS),glutathione peroxidase, haloperoxidase, catalase, cytochrome cperoxidase, horseradish peroxidase, peanut peroxidase, soybeanperoxidase, turnip peroxidase, tobacco peroxidase, tomato peroxidase,barley peroxidase, or peroxidasin. In particular embodiments, theperoxidase is horseradish peroxidase.

In some embodiments, a single FRP enzyme is used. In other embodiments,a combination of FRP enzymes is used, such as any two or threeoxidoreductase enzymes selected from any of the above classes orsub-classes therein. In some embodiments, a combination of FRP enzymes(e.g., EC 1 enzymes) is used. In particular embodiments, a combinationof EC 1.1 enzymes is used. In other particular embodiments, acombination of EC 1.10 enzymes is used. In other particular embodiments,a combination of EC 1.11 enzymes is used. In other embodiments, acombination of any of the particular FRP enzymes described above and aperoxidase is used (e.g., a combination of a EC 1.1 or EC 1.1.3 enzymeand a peroxidase). When a combination of FRP enzymes is used, the two ormore enzymes may be arranged in a core-shell type of arrangement, i.e.,a first FRP enzyme is either in a core portion or surface portion of themagnetic nanoparticle or aggregate thereof, and a second (different) FRPenzyme covers the region where the first FRP enzyme is located. Thesecond FRP enzyme may be an aggregate of the magnetic nanoparticle or onthe surface thereof, overlaying the first enzyme. In the case ofmultiple enzyme systems, manipulating the distribution of the differentenzymes within the mesoporous aggregates offers the advantage ofdecoupling the different reactions and permitting diffusion of thesubstrates and products of the reactions from one layer to another layeror to the core of the BNCs. Therefore, when performing the enzymaticreactions in the confined pore structures of the BNCs, core/shelldistributions offer the possibility of better controlling the kineticsof the different entrapped FRP enzymes. Combining enzymes that performsimilar reactions (such as two, or more, peroxidases or a peroxidase anda laccase for example) but having different reaction requirements(substrates, substrate concentration, etc.) can beneficially increasethe versatility of the BNCs to perform in broad and variable processconditions at a high level of efficiency. Combining enzymes with coupledreactions can ensure the production of the substrate in the vicinity ofthe enzyme and bypass the need for hazardous and labile chemicalsubstrates, such as hydrogen peroxide. For example, a glucose oxidaseenzyme can generate hydrogen peroxide from glucose, which is aninexpensive and non-hazardous compound.

The invention is also directed to methods of producing theenzyme-included (i.e., enzyme-bound, enzyme-trapped, or enzyme-embedded)magnetic nanoparticles and aggregates thereof. In particularembodiments, the enzyme-included magnetic nanoparticles or aggregatesthereof are prepared by including an FRP enzyme in the reactionconditions used for preparing the magnetic nanoparticles or aggregates.For example, an FRP enzyme can be included in the process of generatingmetallic nanoparticles (e.g., cobalt, nickel, or iron) or metal oxidemagnetic nanoparticles (e.g., an oxide of cobalt, nickel, or iron).Synthetic methods for producing metallic and metal oxide magneticnanoparticles are well known in the art. One known method for producingmetallic nanoparticles includes the reduction of metal ions (e.g., as ametal salt) in solution. The reduction can be accomplished by, forexample, a reductive chemical method (e.g., by reaction with a reducingagent, such as hydrogen, a borane, hydrazine, hypophosphate, or citrate)or a reductive or decompositional physical method (e.g., sonication orthermal treatment in solution). The method may alternatively decompose azerovalent metal complex (e.g., a Ni⁰ carbonyl or phosphine complex) by,for example, sonication, thermal treatment, or exposure to a radiativesource, such as ultraviolet light. A particular known method forproducing metal oxide magnetic nanoparticles involves alkaline reactionwith a metal salt (e.g., a metal halide) under conditions where metaloxide nanoparticles precipitate. For example, by well-establishedprocedures, iron oxide nanoparticles can be produced by co-precipitationof iron (II) and iron (III) ions (e.g., as found in FeCl₂ and FeCl₃) insolution by reaction with a base, such as NaOH. The FRP enzyme can beincluded in any such method, as long as the method is not substantiallydetrimental to the activity of the FRP enzyme.

In other embodiments, the magnetic nanoparticles or aggregates thereofare first prepared, and then FRP enzyme is included on or in themagnetic nanoparticles or aggregates thereof. Particularly in thesituation where the magnetic nanoparticles or aggregates are porous, theFRP enzyme can be embedded into the pores of the magnetic nanoparticlesor aggregates in an aqueous-based solution by simple diffusion,adsorption, or self-assembly. In other embodiments, the surfaces and/orpores of the magnetic nanoparticles or aggregates thereof arederivatized with a bonding agent that causes or promotes bonding of theFRP enzyme to the magnetic nanoparticles or aggregates thereof. Thebonding agent can be, for example, a difunctional linker that possessesa reactive end that binds to the magnetic nanoparticle and anotherreactive end that binds to the FRP enzyme. In the case of metallicnanoparticles, the reactive end that binds to the magnetic nanoparticlecan be, for example, an amino, mercapto, mercaptoether, or phosphinegroup. In the case of metal oxide nanoparticles, the reactive end thatbinds to the magnetic nanoparticle can be, for example, a phosphate,phosphonate, sulfate, or sulfonate group. In either case, the reactiveend that binds to the FRP enzyme can be, for example, any of theamine-reactive groups (e.g., N-hydroxysuccinimide group) known in theart, or any of the other groups known in the art for conjugating anenzyme or other protein to another moiety. The bonding agent mayalternatively be, for example, based on affinity coupling, e.g.,producing a magnetic nanoparticle-biotin or -avidin conjugate andreacting this with a FRP-avidin or -biotin conjugate, respectively.

The magnetic nanoparticles or aggregates thereof or BNCs thereof mayalso be coated with a noble metal, such as gold, platinum, or palladium.Any suitable method for coating the magnetic nanoparticles may be used.For example, in particular embodiments, magnetic nanoparticles aredispersed in a solution containing a noble metal salt, and the noblemetal salt subjected to reducing conditions. The foregoing method can befacilitated by binding difunctional molecules onto the surface of themagnetic nanoparticles before the noble metal salt is reduced. Thedifunctional molecules used for this purpose should contain a portionuseful for binding to the magnetic nanoparticles (as described above) aswell as a noble metal binding portion (e.g., an amine, thiol, phosphine,or chelating moiety) for binding noble metal ions. Optionally, oncemetal ions are bound to the nanoparticle surface, the magneticnanoparticles can be washed of excess noble metal salt (e.g., byfiltration or decanting). Since noble metal ions are attached to thesurface, the foregoing methodology provides a more selective method forproducing a noble metal coating (i.e., without concomitant production ofnoble metal nanoparticles) as well as a more uniform coating. In someembodiments, the noble metal coating is applied before FRP enzyme isincluded with the magnetic nanoparticles, in which case FRP enzyme islater bonded to the noble metal coating. The FRP enzyme can be bonded tothe noble metal coating by, for example, functionalizing the noble metalcoating with difunctional molecules that bind to the noble metal coatingand possess another reactive group for binding to the FRP enzyme.

The enzyme-containing magnetic nanoparticles or aggregates thereof, ornoble-metal coated versions thereof, may also be bonded or adhered onto(i.e., be made to reside onto) the surface of ferromagneticmicroparticles. In one embodiment, the enzyme-containing magneticnanoparticles or aggregates thereof, or noble-metal coated versionsthereof, are made to adhere onto the surface of ferromagneticmicroparticles by contacting them in an aqueous-based solution andallowing the nanoparticles to adhere onto the surface of themicroparticles. In other embodiments, the nanoparticles andmicroparticles are suitably functionalized with surface agents tofacilitate a binding interaction, which may be based on, for example,covalent, ionic, affinity, hydrogen bonding, or van der Waals(dispersion) interactions.

In another aspect, the invention is directed to a process fordepolymerizing lignin, i.e., a lignin depolymerization process, in whichany of the enzyme-bound magnetic nanoparticles or aggregates thereof(i.e., BNCs) described above is used for depolymerizing or facilitatingthe depolymerization of lignin. The lignin being depolymerized can beany lignin-containing material. The precursor lignin can be any of awide variety of lignin compositions found in nature or as known in theart.

As known in the art, there is no uniform lignin composition found innature. Lignin is a random polymer that shows significant compositionalvariation between plant species. Many other conditions, such asenvironmental conditions, age, and method of processing, influence thelignin composition. Lignins differ mainly in the ratio of three alcoholunits, i.e., p-coumaryl alcohol, guaiacyl alcohol, and sinapyl alcohol.The polymerization of p-coumaryl alcohol, coniferyl alcohol, and sinapylalcohol forms the p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S)components of the lignin polymer, respectively. The precursor lignin canhave any of a wide variety of relative weight percents (wt %) of H, G,and S components. Besides the natural variation of lignins, there can befurther compositional variation based on the manner in which the ligninhas been processed. For example, the precursor lignin can be a Kraftlignin, sulfite lignin (i.e., lignosulfonate), or a sulfur-free lignin.

Lignin is the most abundant aromatic based biopolymer on Earth, but itis chemically recalcitrant to conversion and bioconversion due to theapparent randomness of its chemical composition and physical structure.Lignin can be considered a “glue” or “epoxy” between polysaccharidefibers that provides strength, rigidity, and protection to the cellwalls of vascular plants. From a chemical standpoint, lignin is a highlyheterogeneous polymer formed by the polymerization of phenyl-propanoidmolecules including coniferyl, sinapyl and coumaryl alcohols via aryllinkages, ether linkages, and carbon-carbon bonds.

Based on the assumption that 100 gallons of ethanol are produced from 1ton of biomass and that biomass (e.g., wood and grass) contains onaverage about 20% lignin, one can quickly estimate that a biorefineryoperating on a 100 million gallon per year capacity would produce about200,000 tons of lignin material. To meet a 20% replacement of gasolinefor the U.S. only by 2020, equivalent to about 35 billion gallons ofethanol, a total of approximately 700 million tons of lignin would beproduced per year. The actual production of lignin, mostly Kraft ligninas byproduct of the paper industry, is approximately 90 million tons peryear worldwide. In other words, the lignin production worldwide would beincreased by more than an order of magnitude.

Lignin can be used for low- or high-priced products based on theapplication and the degree of chemical purity. Until recently, marketsfor lignin products have not been large, competitive, or attractiveenough to compensate for the cost of isolation and purification comparedto the recovered energy derived from its burning. This is mainly becausethe cost of oil is still low enough and the supplies are high enough toprovide the building blocks for the chemical and material industries.However, in a carbohydrate economy framework based on biofuels andbioproducts co-production, high-purity isolated lignin dedicated forconversion could be estimated at $1.10 per kg of raw material comparedto $0.04, when used for co-firing. Low-end applications are mostlydirected to dispersants, soil conditioners for carbon sequestration,adsorbents for fertilizers and pesticides, as well as fuels, whichrequire little or no further conversion after extraction. High-endapplications requiring depolymerization of lignin include the productionof phenolic precursors (DMSO, vanillin, phenol, and aromatic compounds)and polymer components (e.g., epoxy resins, polyurethane foams, phenolicresins powders, carbon fibers and glue and binders).

In nature, the conversion of lignin is performed by specialist microbes,particularly fungi and bacteria. Lignocellulosic bacteria and fungushave the ability to depolymerize lignin in order to gain access tocellulosic fractions of biomass. To that end, lignocellulosic bacteriaand fungus excrete an array of oxidoreductase enzymes, which includelaccases, oxidases, and peroxidases, along with organic acids andH₂O₂-producing catalases. The most potent oxidoreductase enzymes areproduced by a specific group of fungi known as white rot fungi, whichspecialize in lignocellulosic degradation. Various types of fungalperoxidases differ in the nature of their substrates.

Lignin peroxidase (LiP, E.C. 1.11.1.14) catalyzes the oxidative cleavageof C—C bonds in a number of model compounds, and oxidizes benzylalcohols to aldehydes or ketones. Typical reactions catalyzed by ligninperoxidases are Cα-Cα cleavage, Cα oxidation, alkyl aryl cleavage,aromatic ring cleavage, demethylation, hydroxylation and polymerization.Lignin peroxidases are involved in the oxidative breakdown of lignin inwhite-rot basidiomycetes. Lignin peroxidase catalyzes the oxidation ofnon-phenolic aromatic rings into aryl cation radicals by H₂O₂. A typicalexample is the oxidation of veratryl alcohol (3,4-dimethoxybenzylalcohol) into veratryl aldehyde (3,4-dimethoxybenz aldehyde) via theintermediary formation of veratryl cation and benzyl radicals: veratrylalcohol+H₂O₂→veratryl aldehyde+2 H₂O. Manganese peroxidase (MnP; E.C.1.11.1.13) has lower redox potentials (up to 1.1 V) than LiP (up to 1.5V) and catalyzes the Mn-mediated oxidation of lignin and phenoliccompounds. This enzyme catalyzes the oxidation of Mn(II) to Mn(III) byH₂O₂. The highly reactive Mn(III) is stabilized via chelation in thepresence of dicarboxylic acid: 2 Mn(II)+2 H⁺+H₂O₂→2 Mn(III)+2 H₂O. Thepurpose of MnP is to generate small and potent oxidizing agents thatdiffuse into the lignified cell wall and achieve depolymerization oflignin from within. Versatile peroxidase (syn. hybrid peroxidase,manganese-lignin peroxidase: VeP EC 1.11.1.16) is a fairly newligninolytic enzyme, combining catalytic properties of manganeseperoxidase (oxidation of Mn(II)), lignin peroxidase (Mn-independentoxidation of non-phenolic aromatic compounds) and plant peroxidase(oxidation of hydroquinones and substituted phenols). Any one or acombination of the above-mentioned peroxidases may be used in the lignindepolymerization process described herein.

In a first embodiment, the lignin-containing material is a form oflignin partially or substantially separated from other components ofwood (e.g., cellulosic and hemicellulosic components), as is generallyprovided from a pretreatment process of lignocellulosic material, thedetails of which are well known in the art of lignocellulosic processingand conversion. The pretreatment process serves to either separatelignin from other components of the lignin-containing source, or toweaken the bonds between lignin and the other components. As is alsowell known in the art, the lignin may be further isolated by, forexample, extraction. In a second embodiment, the lignin-containingmaterial is a lignin-containing consumable product, such as paper orcardboard, which may or may not be pretreated. In a third embodiment,the lignin-containing material is a lignin-containing natural source(i.e., raw lignocellulosic material), such as woodchips, grasses (e.g.,switchgrass and mixed grasses), corn stover (e.g., leaves, husks,stalks, or cobs of corn plants), sugarcane, saw dust, hemp, or acombination thereof, all of which are generally pretreated to make thelignin sufficiently available for depolymerization.

In the lignin depolymerization process, any of the enzyme-bound magneticnanoparticles or aggregates thereof, described above, is contacted witha lignin-containing material under conditions where partial or completedepolymerization of lignin occurs by free-radical activity of theenzyme-bound magnetic nanoparticles or aggregates thereof. Theenzyme-bound magnetic nanoparticles or aggregates thereof and thelignin-containing material are generally made to contact by combiningthem in an aqueous solution, such as an aqueous solution used in apretreatment process of the lignin-containing material. In someembodiments, a room temperature condition (e.g., at least 15, 18, 20, or22° C. and up to 25° C. or 30° C.) is used during the depolymerizationprocess. In other embodiments, an elevated temperature condition (e.g.,above 30° C., or at least or above 35, 40, 45, 50, or 60° C., or up tothe temperature that the FRP enzyme degrades or suffers a substantialloss in activity) is used during the depolymerization process. In otherembodiments, a reduced temperature condition (e.g., below 15° C., or upto or below 10, 5, or 0° C.) is used during the depolymerizationprocess. By being depolymerized, the lignin is broken down into shortersegments compared to its original form. A complete depolymerizationresults in the conversion of all or a substantial portion (e.g., atleast 80, 90, or 95%) of the lignin into at least one or more of thebasic building blocks of lignin, i.e., coniferyl, sinapyl, and coumarylalcohols, and derivatives thereof. A partial depolymerization generallyresults in less than 80%, or up to 70, 60, 50, 40, 30, 20, 10, 5, or 1%of lignin being converted to primary building blocks, with the rest ofthe lignin being converted to segments containing two, three, four, or ahigher multiplicity (even up to 10, 20, 50, 100, 200, 500, or 1000) ofbuilding blocks (e.g., p-hydroxyphenyl, guaiacyl, and syringyl unitsderived from coumaryl, coniferyl, and sinapyl alcohols, respectively).Since different degrees of lignin depolymerization may be preferred fordifferent applications, the depolymerization conditions can be suitablyadjusted to provide an appropriate degree of depolymerization or tofavor one or more types of depolymerization products over others.

Since each lignin-containing material has a different distribution andrelative amount of each building block, the relative amount of eachproduct produced from depolymerization is very much dependent on thetype of lignin-containing material. Other depolymerization products,e.g., aromatic aldehydes, ketones, alcohols, and acids, are generallyalso produced during the polymerization process, typically in lesseramounts. In embodiments where such other products are not desired, theymay be advantageously minimized or eliminated as a product by adjustmentof reaction conditions, including appropriate selection of the FRP-boundmagnetic nanoparticle or aggregate thereof.

Any of the enzyme-bound magnetic nanoparticle and aggregate compositionsdescribed above can be used for the lignin depolymerization process. Inparticular embodiments, the FRP enzyme used in the lignindepolymerization process is a peroxidase, and particularly, alignin-degrading peroxidase, such as a lignin peroxidase, versatileperoxidase, manganese peroxidase, or combination thereof (including acore-shell combination thereof). The FRP enzyme may also moreparticularly be a fungal, microbial, or plant peroxidase. In specificembodiments, the FRP enzyme is a system of two FRP enzymes, such as afungal peroxidase combined with a glucose oxidase, or a peroxidaseand/or oxidase combined with a laccase.

In some embodiments, the lignin depolymerization process is coupled(i.e., integrated) with a downstream process in which depolymerizationproduct produced in the lignin depolymerization process is used for theproduction of other products. The downstream process may convert lignindepolymerization product into, for example, biofuel or an industrialchemical product, e.g., a polymer, plastic, polymer precursor (monomer),solvent, adhesive, paint, detergent, lubricant, food product, medicinalproduct, or aroma, or a precursor therefore. The downstream process mayalternatively incorporate the lignin depolymerization product into anysuch end product.

In some embodiments, the lignin depolymerization process is coupled withan upstream process in which lignin-containing material is provided foruse in the lignin depolymerization process described herein. Theupstream process can be, for example, a paper or pulp producing process,a biomass-to-biofuel process (i.e., where primarily cellulosic materialis hydrolyzed and converted to biofuel), or a biomass-to-ethanolfermentation process (i.e., where primarily cellulosic material ishydrolyzed and converted to ethanol).

In another aspect, the invention is directed to a process for removingaromatic contaminants from water (i.e., a water remediation process). Inthe process, water contaminated with one or more aromatic substances iscontacted with any of the enzyme-bound magnetic nanoparticles oraggregates thereof, described above, to cause the aromatic substances toprecipitate, i.e., as insoluble material. The precipitated (i.e.,sedimented) material is preferably then further separated, such as bycentrifugation or settling, and removed from the water by, for example,filtration or decanting. Without being bound by any theory, it isbelieved that the aromatic substances react with free radicals producedby the enzyme-bound magnetic nanoparticles to produce a polymerizedmaterial derived from the aromatic substances. The aromatic contaminantcan be any aromatic substance, including those more commonly found incontaminated water. In some embodiments, the aromatic contaminant isbenzene, or a benzene derivative, such as a halogenated benzene (e.g.,chlorobenzene, dichlorobenzenes, bromobenzenes, or a polychlorinatedbiphenyl, i.e., PCB), alkylbenzene (e.g., toluene, ethylbenzene, or axylene), phenolic substance (e.g., phenol, resorcinol, catechol, or acresol), etherified benzene (e.g., anisole), fused ring compound (e.g.,naphthalene, or polyaromatic hydrocarbon), aniline substance (e.g.,aniline and N-alkyl or N,N-dialkyl substituted anilines), or benzoicacid compound (e.g., benzoic acid, esters thereof, andhydroxy-substituted derivatives of benzoic acid). In other embodiments,the aromatic contaminant is a heteroaromatic substance, such as furan,pyran, dioxin, thiophene, pyridine, pyrazine, pyrimidine, pyrrole,imidazole, indole, and derivatives thereof.

Any of the enzyme-bound magnetic nanoparticle and aggregate compositionsdescribed above can be used for the water remediation process. Inparticular embodiments, the FRP enzyme used in the water remediationprocess is horseradish peroxidase, or horseradish peroxidase incombination with an oxidase.

In another aspect, the invention is directed to a process forpolymerizing monomers polymerizable by a free-radical mechanism. In theprocess, one or more types of monomers are reacted with any of theenzyme-bound magnetic nanoparticles or aggregates thereof, describedabove, to cause the monomers to polymerize. The monomers can be, forexample, any of the substances provided above for the water remediationprocess. In particular embodiments, the monomers are or includevinyl-addition monomers. Upon polymerization, a vinyl-addition polymeris produced. Some examples of such monomers include ethylene, propylene,butadiene, the acrylates and esters thereof, methacrylates and estersthereof, acrylonitriles, vinyl acetate, styrene, divinylbenzene, vinylfluorides, and vinyl chlorides. In other embodiments, the monomers arephenolic compounds. Upon polymerization, a phenolic resin or polymer isproduced. The polymerization process can utilize any of the conditionsand apparatuses well known in the art for practicing polymerizationreactions, and in particular, free-radical initiated polymerizationreactions.

For any of the processes described above, the enzyme-bound magneticnanoparticles or aggregates thereof can advantageously be captured bymagnetic separation in order to prevent contamination of the finalproduct. Moreover, a further advantage of the enzyme-bound magneticnanoparticles or aggregates thereof described herein is their ability inmany cases to retain their activity and re-form after capture, whichpermits them to be re-used after capture. BNCs showing a loss ofactivity after several cycles can advantageously be easily extracted andconcentrated to their solid form to provide a less wasteful and moreefficient process. Metal-coated BNCs can be repurposed by denaturationof the enzymes, sonication, and purification in order to be restored andre-used with fresh functional enzymes. Micro-Bionanocatalysts(Micro-BNC) made of self-assembled BNCs magnetically trapped on asurface are attractive for process applications that use lower intensitymagnetic fields. Larger and denser ferromagnetic particles have a highermass susceptibility compared to mesoporous and low density aggregates ofMNPs. MicroBNCs maintain stable, nanosized, and mesoporous structures,which helps to maintain enzyme activity while increasing the overalldensity and mass susceptibility of the magnetic catalyst. Theseultrastructures lend themselves to easier manipulation by externalmagnetic fields as produced by permanent small magnets and weak fieldelectromagnets. The reaction solution can be purged and replaced whilethe Micro-BNCs are magnetically trapped, hence allowing for sequentialuse of the Micro-BNC as long as the enzyme retains process levelactivities.

Examples have been set forth below for the purpose of illustration andto describe the best mode of the invention at the present time. However,the scope of this invention is not to be in any way limited by theexamples set forth herein.

EXAMPLES

Reagents and Instrumentation

The enzymes used in this study were Horseradish Peroxidase (HRP, E.C.1.11.1.7, type VI-A), Lignin Peroxidase (LiP, E.C. 1.11.1.14), ManganesePeroxidase (MnP; E.C. 1.11.1.13) Versatile Peroxidase (syn. hybridperoxidase, manganese-lignin peroxidase: VeP EC 1.11.1.16), and laccase.All enzymes used herein were obtained from commercial sources.Horseradish Peroxidase (HRP, E.C. 1.11.1.7, type VI-A) had aReinheitszahl index (OD A₄₀₃/A₂₈₀) of around 2.9. High activity formswere obtained by further purification by FPLC (AKTA Explorer, GEBioscience) using an anionic exchange column (Resource Q, GEBioscience). Laccase was used without further purification. Proteinsignal was monitored at 280 nm and the heme signal at 405 nm. For eachenzyme, the fractions with a RZ above 1 were pulled together,concentrated and aliquoted. Phenol, homovanillic acid, veratryl alcohol,methoxylphenol, 4-aminoantipyrine (AAP), glucose (the foregoing being at98% purity), sodium phosphate buffer (PBS) of pH 7.4 and 67 mM,magnetite microsphere powder, manganese sulfate, hydrogen peroxide,FeCl₃.6H₂O, and FeCl₂.4H₂O, O-phosphoethydiamine, goldtetrachlorohydrate, and malonate and tartrate disodium salts wereobtained from commercial sources.

Synthesis of Iron Oxide Magnetic Nanoparticles

Magnetite nanoparticles were synthesized by co-precipitation of Fe²⁺ andFe³⁺ under alkaline conditions in a bubbling nitrogen atmosphere at 25°C. (M25) or 90° C. (M90). An acidic solution (25 ml) of the iron salts(2 g of FeCl₂.4H₂O and 5.2 g of FeCl₃.6H₂O) was added dropwise to NaOH(250 ml, 1.5 M) under constant stirring. Non-oxidizing conditions wereachieved by bubbling all solutions with nitrogen for 15 minutes prior toreaction. The instantaneous black precipitation of Fe₃O₄ was capturedwith a neodymium magnet, washed, and neutralized, and kept in distilledwater until further use.

Coating of Iron Oxide Magnetic Nanoparticles with Gold

The coating of magnetite nanoparticles was achieved by mild reduction ofgold tetrachloroaurate ions onto O-phosphoethydiamine (OPEA)functionalized MNPs under sonication. The coating procedures wereperformed under nitrogen using a modified rotary evaporator apparatuscoupled to a sonic bath. Briefly, 20 mg of MNPs (1 mg/ml) were sonicatedfor 30 min. OPEA (2 g) was added and allowed to react with the MNPs (40mL final volume) under sonication and rotation for 2 hours.OPEA-functionalized MNPs were capture with a rare-earth magnet andrinsed 3 times to remove excess OPEA. OPEA-functionalized MNPs werere-suspended in milliQ water with 100 μL of nitric acid (1N) and wereagitated for 1H under sonication. Gold tetrachloroaurate (10 up to 30mg) was added to the OPEA-functionalized MNPs and allowed to react for30 minutes. The temperature of the sonic bath was raised to 85° C., atwhich point citric acid/citrate (50:50, 50 mM total, pH5.5) wasinjected. The gold coating was performed under sonication and fastrotation speed in varying volume (up to 100 mL) and different coatingtimes (up to 60 min). The reaction vessel was placed on ice and thereaction was stopped by adding 5 g of CaCO₃. MNP-OPEA-Au nanoparticleswere captured magnetically, rinsed four times to remove excess reagents,and stored until further use.

Bionanocatalyst (BNC) Synthesis

All BNCs were formed with either M90 or M25 magnetite nanoparticles,gold-coated or not. The BNCs were formed with either Horseradishperoxidase, Lignin Peroxidase, Manganese Peroxidase or VersatilePeroxidase. Typical BNC synthesis required the MNPs to be initiallymonodispersed (i.e., individualized and non-aggregated MNPs). MNPs weresonicated in an ultrasonic bath for 20 minutes at room temperature andused immediately to form the BNCs. In preliminary experiments, theeffects of buffer composition, buffer strength, and incubation time wereinvestigated to measure their influence on the formation of the BNCs andthe activity of the enzymes. Formation of BNCs in water for more thanone hour was found to be the most efficient due to the lack of ioniccompensative charges; this protocol was implemented for the rest of thestudy. The final concentration of the peroxidases used for the assaysranged from 0.1 to 10 nM. It was also found that the increased activitywas higher for BNCs with 50% or less saturation (per surface area), andtherefore, the quantity of MNPs added to the enzymes were adjustedaccordingly. Typically, the final concentrations of MNPs in the assaywere between 0.5 and 50 μg·ml⁻¹. A typical ratio is 1 nM of enzyme (HRP)for 2 μg·ml⁻¹ of MNPs (final concentrations in the assay). The enzymeand the MNPs were added simultaneously and were incubated under constantagitation at 4° C. for at least 12 hours. Typically, the BNCs wereformed in stock solution at 5 or 10 times the concentration needed toperform the biochemical reactions. BNCs were finally diluted to thefinal concentration required in the assay just before the assay. For thesynthesis of core/shell poly-enzyme systems BNCs, a core BNC was firstformed by reacting the first enzyme (peroxidase or glucose oxidase coreenzyme) and sonicated MNPs for 7 hours, then the second enzyme(peroxidase or glucose oxidase=shell enzyme) was added with theappropriate ratio of sonicated MNPs and incubated for a least 7 hours.For random BNCs of glucose oxidase and peroxidase, the sonicated MNPsand enzymes were added simultaneously and incubated for 14 hours.

Micro-Bionanoparticle (Micro-BNC) Synthesis

Micro-BNCs (μBNCs) were synthesized by reacting formed BNCs withcommercial submicrometric magnetite particles. Submicroparticles weresuspended in water and sonicated for 20 minutes in an ultrasonic bath.Preformed BNCs and sonicated microparticles were incubated for 6 hoursin water under constant agitation and at 4° C. The quantity ofsubmicrometric commercial particles was at least one order of magnitudehigher than the quantity of the MPNs forming BNCs on a weight basis. Thecapture of the BNCs was considered complete when no nanoparticle wasdetected in the supernatant after magnetic capture with a small magnet.

Characterization of Magnetic Nanoparticles and Mesoporous Aggregates

Magnetic properties were measured using a MPMS XL® (Quantum Design)magnetometer utilizing Superconducting Quantum Interference Device(SQUID) technology. Magnetization hysteresis curves were determined at300K for external magnetic fields ranging from 40 Oe to 50 kOe.Transmission electron microscopy (TEM) image processing was used tomeasure the average particle size, size distribution and cluster sizes.TEM measurements were performed with an UHV-STEM microscope (VG, UK).Images were processed using Image Analysis Image J software (NIH,Washington D.C.) and JMicroVision (V1.27). Nanoparticle and cluster sizedistributions were calculated from a minimum of 1000 particles. Nitrogenadsorption-desorption isotherms were obtained on a Micrometrics ASAP2020 physisorption instrument. Pore size distributions were calculatedfrom the N₂ adsorption isotherm using the Barrett-Joyner-Halenda (BJH)method.

Elucidation of Magnetic Nanoparticles

The X-ray diffraction pattern shown in FIG. 1 confirms that the MNPs aremade of magnetite. The synthesis described above uses low-cost reagentsand can be easily tailored in size (FIGS. 2A, 2B) and magnetic field(FIG. 3). As shown by FIGS. 2A and 2B, M90 had a more uniform sizedistribution with an average size of 10 nm (±1), while M25 had a broadersize range with an average size of 8 nm with particles as small as 5 nm.As shown by FIG. 3, both MNPs had overall negligible remanentmagnetization (M_(R)), which is indicative of superparamagnetism. Asshown by FIGS. 4A-4C and 5A, at the concentration of magnetite used forthe assays, M25 MNPs were mostly monodisperse with small clusters andfree nanoparticles, whereas M90 formed large clusters of 100 nm indiameter on average. As shown by FIGS. 7 and 8, the total pore volumewas 0.39 cm³·g⁻¹ and 0.14 cm³·g⁻¹ for M90 and M25, respectively. Theaverage density of the clusters when corrected for the porosity wasfound to be ρ_(M25) of 3.03 g·cm⁻³ and ρ_(M90) 1.72 g·cm⁻³.

Quantification of Entrapped Enzyme Using Horseradish Peroxidase (HRP)

Quantification of denatured HRP with magnetite nanoparticles wasachieved by using a high-throughput FTIR spectrometer (HTS-XT-Vertex70,Bruker, Germany). Fifty microliters of BNC slurry was dried out on atransmittance silicon 96-well plate at 60° C. under vacuum for one hourand left to dry under vacuum at room temperature overnight. The spectrawere recorded between 4,000 and 400 cm⁻¹, 32 scans, and the backgroundwas recorded before each sample. Samples were analyzed in triplicates;on-plate standards of magnetite and magnetite plus HRP were used tomeasure the concentration of proteins and nanoparticles in each sample.Adsorption isotherms parameters were extracted by fitting the quadraticform of the Langmuir equation using a least-square fitting method usingMatlab software:

$\begin{matrix}{Q = \frac{Q_{m}K_{a}C^{*}}{1 + {K_{a}C^{*}}}} & (1)\end{matrix}$

In Formula (1) above, Q is the adsorbed enzyme on the surface(nmol·m⁻²), C* is the initial enzyme concentration (nmol·m⁻²), Q_(m) isthe maximum amount of bound enzyme (nmol·m⁻²), and K_(a) is theadsorption constant (m²·mol⁻¹). These parameters were applied in thekinetic experiments to directly calculate the bound fraction of HRP atequilibrium from the initial concentration of HRP.

Characterization of Peroxidase Enzyme Activities

Phenol/AAP assay: The peroxidase activity of native HRP and BNC wasmonitored using the chromogenic phenol/AAP assay that generates phenoxyradicals that readily react with aminopyrene to form the pink-coloredquinoneimine dye. An automated plate-reader (Synergy 4, Biotek) withinjection capabilities and temperature-controlled chamber was used torecord the absorbance of the solution at 510 nm in 96-well plates (4replicates) for 30 minutes. The standard reagent concentrations of theassay (200 μl) were 80 mM and 13 mM for phenol and AAP respectively.Different buffer and buffer strength were tested. Hydrogen peroxide wasinjected to initiate the reaction with concentrations ranging between10⁻⁷ M and 1 M. The background contribution due to the nanoparticles andsubstrates was subtracted. For HRP, the quantity of free enzymes wascalculated by difference with the bound amount estimated with theLangmuir adsorption parameters as the quantity of free enzyme could notbe estimated directly because of low HRP concentrations and highbackground from the MNPs. For each run, a velocity standard curve wasestablished for the free enzyme and used to correct for the contributionof the free enzyme to the total activity when not all enzyme moleculeswere bound. The velocities (V) and specific activity, A(mmol_(product)·s⁻¹·mmol_(enzyme) ⁻¹) were calculated based on theinitial rates of the reaction.

Fungal Peroxidase activities using model substrates: Colorimetric assayswere performed in 96-well UV transparent microplates (Falcon) usingstandardized colorimetric protocols in sodium tartrate or malonate (LiP:Veratryl alcohol, pH 3, 310 nm; MnP: 2,6 dimethoxyphenol, pH 4.5, 468 nmand 270 nm for the formation of Mn³⁺-organic acid complex; VeP: veratrylalcohol or 2,6 methoxyphenol, pH 4.5, 310 nm, 468 nm, 270 nm).

Peroxidase activities with Homovanillic acid (HVA) fluorescent assay: Afluorescent assay was developed to measure the initial rate of theperoxidase kinetics. HVA free radicals can polymerize to form afluorescent dimer (λ_(ex) 310 nm, λ_(em) 405 nm). An automatedsequential procedure was implemented on a Biotek Plate Reader withsyringe mixing capabilities to perform the dilution of the reactioninitiators (H₂O₂ or glucose). A chemically-reacted stock of fluorescentdimer was synthesized by reacting HVA (10 mM) with sodium ferrocyanide(15 mM) and ammonium hydroxide (15 mM) and was used as quantitativestandards in the buffers used for the enzymatic reactions. All thereactions were performed in triplicate and data expressed in relativefluorescent units (mRFU).

Lignin depolymerization assays: A Kraft lignin depolymerization assaywas conducted using HRP, LiP, VeP and MnP, and BNCs thereof. MnP and VePassays were performed in presence of manganese. Assays were performed intriplicates in sodium tartrate or sodium malonate buffer, pH 5.5. Kraftlignin slurries (10 mg/ml) were incubated for 1 or 4 hours then filteredthrough 0.2 μm pore membrane filter to remove particulates. The UV-Visspectra of the solutions were acquired with a Biotek Plate Reader. Therelease of aromatic molecules from lignin depolymerization was monitoredat 280 and 310 nm. The spectra were corrected for the background.

Phenol polymerization assays: The phenol removal assay was a two-stepprocedure. The first step consisted in forming polyphenols withHorseradish Peroxidase in 1 mM PBS buffer. Reaction volumes were fixedat 2 ml or 10 mL. The final concentration of phenol was fixed at 1 mMand HRP at 30 nM. The BNCs formed with M90 were varying inenzyme-to-nanoparticles ratio. The second step was the precipitation ofthese polyphenols by adding sodium chloride (500 mM). The samples werecentrifuged at 12,000 g for 20 minutes and the supernatant collected.The soluble phenol in solution was measured at 280 nm with a BiotekPlate Reader.

Kinetic Parameters

The specific activity, A (mmol_(product)·s⁻¹·mmol_(HRP) ⁻1), wascalculated as the ratio V/mmol_(HRP) using the extinction of therespective products formed at the wavelength monitored. A H₂O₂ substrateinhibition model derived from the ping-pong bi-bi 2 substrate inhibitionmodel was used to extract the kinetic parameter of the reaction with aleast-square fitting method using GraphPad Prism (La Jolla, Calif.,USA). The modified equation from the model is:

$\begin{matrix}{V = \frac{V_{\max}\left\lbrack {H_{2}O_{2}} \right\rbrack}{K_{m} + {\left\lbrack {H_{2}O_{2}} \right\rbrack\left( {1 + {\left\lbrack {H_{2}O_{2}} \right\rbrack/K_{i}}} \right)}}} & (2)\end{matrix}$

In Formula (2), V_(max) is the maximum enzyme velocity (mmol·s⁻¹), themaximum rate the enzyme reaction can achieve, expressed in the sameunits as V, K_(m) is the Michaelis-Menten constant (mM), K_(i) is theinhibition constant for H₂O₂ (mM). k_(cat) (s⁻¹) was calculated fromV_(max) and the total quantity of bound HRP.

Formation, Characterization and Activities of Magnetite BNCs Formed withHRP

The entrapping of enzyme molecules in the MNP clusters was confirmed bythe overall increase in size of the BNC clusters (FIGS. 4A and 5G). BothM25 MNPs (FIG. 4 B) and M25-BCNs (FIG. 4 C) had small cluster sizesalthough M25-BNCs size was slightly increased and had bigger clusters. Acomparison of M90 MNPs (FIG. 5A) and M90-BNCs (FIG. 5D) demonstratesthat the increase in size of the nanoparticle aggregates was higher forM90 than for M25. Moreover, the aggregates of M90 BNCs (FIGS. 5E and 5F)had a higher density, i.e. made of more MNPS and more compacted MNPs,than the aggregates of M90 alone (FIGS. 5B and 5C). The entrapping ofenzyme molecules in the MNP clusters was further confirmed by thedifference in HRP adsorption behavior between M25 and M90 (FIG. 6). BothBNCs were mesoporous (FIGS. 7 and 8) with pores below 50 nm. M90-BNCshad a higher total pore volume and higher average pore size thanM25-BNCs. The differences in the formation of the complexes wasconsequently attributed to the ultrastructure of the MNPs clustersresulting from differences in magnetization of the nanoparticles andresulting mesoporosity of the clusters. In particular, M90-BNCself-assembly appears to result from a dual mechanism of surfaceadsorption and molecular entrapment in the mesoporous aggregates.

M90 had a higher K_(a), thus indicating a higher affinity for the enzymecompared to M25 (FIG. 6). In quantifying the entrapped enzymes, it wasfound that less than 50% of the total HRP loaded could bind to M25 toform the M25-BNC complex when 1 nM HRP and 4 μg·ml⁻¹ magnetite particleswere used, while 100% of the enzyme was captured in the case of M90. Asshown by FIGS. 11A, 11B, 12A, 12B, and 13, the increased in activitiesvaried as the BNCs were formed in different conditions. The most activeBNCs were formed in ultrapure water at pH 6.5, which implies thepresence of Fe—OH²⁺ cationic species on the nanoparticle surfaces. Atthis pH and in the absence of any other compensating charges, a largesurface of the HRP molecule remains negatively charged, which can allowfor the formation of complexes with magnetite via electrostaticinteractions. When the BNCs formed in PBS were tested (FIG. 11A), theincrease in activity was lower than for the BNCs formed in water (FIG.11B). BNCs formed in PBS had the lowest increase in activity of all.When the BNCs formed in malonate were tested (FIG. 12A), the increase inactivity was in the same range or lower than for the BNCs formed inwater (FIG. 11B). When the BNCs formed in tartrate were tested (FIG.13A), the increase in activity was lower than for the BNCs formed inwater (FIG. 13B). As also shown by FIGS. 11B, 12B and-13B, only BNCsformed in water consistently showed a higher increase in activity thanBNCs formed in other ionic buffers, even at higher concentration ofsalts, when used in other buffers for the assays.

Temporal free-radical concentrations were measured using the phenol/AAPassay, and these measurements were used to calculate the normalizedactivities of the BNC. Normalized activities were calculated as theratio of the BNC activity divided by the free enzyme activity at thesame concentration. For M90-BNC, the maximum activity compared to thefree enzyme was reached after 1 hour of pre-incubation. The increase ofnormalized activity was shown to be stable over 24 hours. As shown byFIGS. 9A and 9B, M25-BNC reached the maximum increased activity after 10minutes of incubation. These results clearly demonstrate that theincreased activity is only due to the immobilized enzyme and not justthe presence of the MNPs in solution. Moreover, as shown by FIG. 10, theresults also demonstrate that the association of HRP with magnetitenanoparticles of different size, magnetism, and ratio yield differentcomplexes with different specific activity. As shown by FIG. 14, thesaturation of the BNCs with the maximal amount of entrapped enzymeconsistently resulted in a decrease in activities.

Initial reaction velocities were used to estimate K_(m), V_(max), K_(i)and k_(cat), as further demonstrated by the plots shown in FIGS. 15-18,respectively. The V_(max) of M25 or M90 MNPs was several orders ofmagnitude lower than those of the free HRP and BNCs. The turnover rates,k_(cat), were very consistent with free HRP, M25-BNC, and M90-BNCdatasets. BNC formed with M25 and M90 both had a higher K_(m). M25-BNChad V_(max), and k_(cat) two to three times greater than the free enzymeat the same concentration. Also, the k_(cat) of M25-BNCs increased withthe fraction of bound enzyme while the K_(i) was in the same range thanthe free HRP. At 0.5 nM of bound enzyme, the efficiencies of the MNPs,as estimated by k_(cat)/K_(m), were 6.75×10³ and 5.5×10³ s⁻¹mM⁻¹ forM25-BNCs and M90-BNCs, respectively. These kinetic results areconsistent with the trends observed with the reaction velocities.M90-BNC had a K_(i) about 10 times greater than the free HRP, while itsV_(max) was similar. The higher K_(i) for M90 indicated the lower extentof substrate inhibition from H₂O₂ compared to the free enzyme andM25-BNCs.

The M90-BNC activities were further investigated for different buffer,pH, and temperature (FIGS. 19A, 19B, 20A, and 20B). A similar increasein M90-BNC activity was also observed in inorganic and organic buffersand across the range of temperature tested. The BNCs were found to bemore efficient at lower temperatures than the HRP alone, as the freeenzyme had higher velocities at higher temperature. The M90-BNCS werefound to be pH sensitive, with the velocities increasing with pH, whilethe normalized activities were pH dependent in inorganic buffers. Nosignificant effect of pH was observed with the organic acid buffers.

BNCs Activities for other Enzyme Systems

Activities of Versatile Peroxidase-BNCs: BNCs were formed with VersatilePeroxidase enzyme (FIG. 21). As shown by the activity plot in FIG. 21,the maximal activities were observed for BNCs formed with M90. Surfacemodifications with organic polymers only resulted in a modest increasein activities but lower inhibition from hydrogen peroxide.

Activities of Manganese Peroxidase-BNCs: BNCs were formed with manganeseperoxidase (FIG. 22). As shown by the velocities plot in FIG. 22,gold-coated MNPs further increase the velocity of the reaction comparedto the non-coated magnetite nanoparticles. Differences were observedbetween the coating conditions (and nanoparticles sizes), resulting indifferent increases in velocities.

Activities of Laccase-BNCs: BNCs were formed with laccase (FIG. 23). Asshown by the assay plot in FIG. 23, the increase in laccase activity wasobserved when the enzyme was used in non-optimal condition (low or highpH). The increase, in activity was not as high as the peroxidase enzymesbut still significant compared to the free laccases. It is noteworthythat laccases do not require peroxide to function, and therefore, arenot subjected to strong substrate inhibition. As previously observed,BNCs provide a strong protection against substrate inhibition and alsoharsher reaction conditions (e.g., pH, temperature, and ionic strength).

Activities of Glucose Oxidase and Peroxidase system: BNCs were formedwith a dual glucose oxidase/peroxidase system (FIGS. 24A-24D and 25). Inthis configuration, the peroxide is provided by the activities of theglucose oxidase in order to activate the peroxidase. A differentconfiguration was tested using the highly monodispersed gold coatedMNPs. As shown by FIGS. 24A-24D and 25, in the ratio of Gox to HRP used,no inhibition of the enzyme system was observed. A core/shell design wasimplemented for which the HRP is at the core of the clusters and the Goxin the outer layers, and vice versa, or randomly distributed. Core/shellBNCs showed increased activities compared to the free enzymes and therandomly distributed BNCs at lower concentration of glucose.Significantly, this system permits replacing hydrogen peroxide withglucose, and glucose is much less costly, more plentiful, and lesshazardous than hydrogen peroxide.

These new BNCs were further immobilized on larger ferromagneticmagnetite submicrometric particles. Scanning electron microscope (SEM)micrographs of commercial magnetic particles and their surfacefunctionalization with BNCs are shown in FIGS. 26A and 26B. Although theaggregates of these ferromagnetic particles are micrometric in size, theindividual crystallites are submicrometric. The ferromagneticmicroparticles readily capture the smaller BNCs that form small clusterson their surface. Although MNPs are fairly magnetic, they require highfield magnets to be captured. Also, the smaller and more monodispersethe MNPs, the longer it takes to capture them with an external magneticfield. The BNC-microparticle clusters substantially overcome thislimitation. Moreover, as shown by FIGS. 27 and 28A-28C, the enzymaticactivities in the BNC-microparticle clusters are similar to the activityof the BNCs (FIG. 27) and can be captured to be re-used or stirred(FIGS. 28A-28C) with very small magnets. This μBNC configuration makesthe catalysts process-ready for real-world applications.

Lignin Depolymerization by Magnetite BNCs Formed with Fungal Peroxidases

Lignin depolymerization using BNCs was demonstrated. A lignindepolymerization assay was conducted in order to detect the productionof soluble aromatics (e.g., coniferyl, sinapyl, and coumaryl alcohols orderivatives thereof). As shown by the absorbance plot in FIG. 29,increased signals were observed at characteristic wavelengths with thefungal peroxidase system. As also shown by FIG. 29, the BNCs formed withhorseradish peroxidase (plant peroxidase) did not release any aromaticmolecules from lignin. As shown by FIGS. 30-33, depolymerization oflignin was observed with fungal peroxidase BNCs.

Polymerization of Phenol with BNCs Formed with HRP

Phenol removal using Horseradish Peroxidase using BNCs was demonstrated.As pictographically shown in FIG. 34, phenol polymerization assays wereconducted using a two-step process: (i) enzymatic polymerization topolyphenols, and (ii) condensation of the polyphenol polymers by sodiumchloride. As shown by the activity plot in FIG. 35, the BNCs formed withHRP and M90 MNPs were more efficient at removing the phenol than thefree enzyme. The foregoing result demonstrates a marked improvement inphenol removal when using BNCs rather than a free HRP system. As shownin FIG. 36, besides the increased extent of phenol removal, there is aH₂O₂ concentration shift to reach the maximum removal (orpolymerization) in the BNC system. This indicates that BNCs have a lowerinhibition from H₂O₂ compared to the free enzyme. Moreover, the BNCsystem offers a broader H₂O₂ concentration range. These featuresdemonstrate that BNCs can be used in unstable and harsher processconditions than their free counterpart.

While there have been shown and described what are at present consideredthe preferred embodiments of the invention, those skilled in the art maymake various changes and modifications which remain within the scope ofthe invention defined by the appended claims.

What is claimed is:
 1. A composition comprising self-assembledmesoporous aggregates of magnetic nanoparticles and a first free-radicalproducing enzyme, wherein said first enzyme is magnetically-entrappedwithout a bonding agent in mesopores formed by said aggregates ofmagnetic nanoparticles and said first enzyme functions by converting adiffusible substrate into a diffusible product.
 2. The composition ofclaim 1, wherein said mesoporous aggregates of magnetic nanoparticleshave an iron oxide composition.
 3. The composition of claim 1, whereinsaid mesoporous aggregates of magnetic nanoparticles have a magneticnanoparticle size distribution in which at least 90% of magneticnanoparticles have a size of at least 3 nm and up to 30 nm, and anaggregated particle size distribution in which at least 90% of saidmesoporous aggregates of magnetic nanoparticles have a size of at least10 nm and up to 500 nm.
 4. The composition of claim 1, wherein saidmesoporous aggregates of magnetic nanoparticles possess a saturatedmagnetization of at least 10 emu/g.
 5. The composition of claim 1,wherein said mesoporous aggregates of magnetic nanoparticles possess aremanent magnetization up to 5 emu/g.
 6. The composition of claim 1,wherein said free-radical-producing enzyme is contained in saidmesoporous aggregates of magnetic nanoparticles in up to 100% ofsaturation capacity.
 7. The composition of claim 1, wherein said firstfree-radical-producing enzyme is an EC 1.11 oxidoreductase.
 8. Thecomposition of claim 7, wherein said free-radical-producing enzyme iscomprised of a peroxidase.
 9. The composition of claim 8, wherein saidperoxidase is a class III peroxidase.
 10. The composition of claim 9,wherein said class III peroxidase is horseradish peroxidase.
 11. Thecomposition of claim 1, wherein said mesopores are characterized by apore size distribution in which at least 90% of the pore volume isattributed to pores having a pore size of at least 2 nm and up to 20 nm.12. The composition of claim 1, further comprising a second enzyme thatis an EC 1.1.3 enzyme, and wherein said first enzyme is a peroxidase.13. The composition of claim 12, wherein said EC 1.1.3 enzyme is glucoseoxidase EC 1.1.3.4.
 14. The composition of claim 12, wherein saidperoxidase is horseradish peroxidase.
 15. The composition of claim 1,wherein said mesoporous aggregates of magnetic nanoparticles andfree-radical producing enzyme further comprise a metallic surface layer.16. Microparticles comprising the composition of claim
 1. 17. Themicroparticles of claim 16, wherein said microparticles comprisemagnetite.
 18. The microparticles of claim 16, wherein saidmicroparticles are ferromagnetic submicrometric particles having a sizeof at least 20 nanometers.
 19. The composition of claim 1, wherein saidmesoporous aggregates of magnetic nanoparticles have a magneticnanoparticle size distribution in which at least 50% of magneticnanoparticles have a size of at least 1 nm and up to 50 nm, and anaggregated particle size distribution in which at least 90% of saidmesoporous aggregates of magnetic nanoparticles have a size of at least10 nm and up to 500 nm, and wherein said free-radical producing enzymeis an oxidoreductase belonging to the EC1 family of enzymes.